This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Formula display:

Abstract

Lactobacillus panis strain PM1 is an obligatory heterofermentative and aerotolerant microorganism that
also produces 1,3-propanediol from glycerol. This study investigated the metabolic
responses of L. panis PM1 to oxidative stress under aerobic conditions. Growth under aerobic culture triggered
an early entrance of L. panis PM1 into the stationary phase along with marked changes in end-product profiles.
A ten-fold higher concentration of hydrogen peroxide was accumulated during aerobic
culture compared to microaerobic culture. This H2O2 level was sufficient for the complete inhibition of L. panis PM1 cell growth, along with a significant reduction in end-products typically found
during anaerobic growth. In silico analysis revealed that L. panis possessed two genes for NADH oxidase and NADH peroxidase, but their expression levels
were not significantly affected by the presence of oxygen. Specific activities for
these two enzymes were observed in crude extracts from L. panis PM1. Enzyme assays demonstrated that the majority of the H2O2 in the culture media was the product of NADH: H2O2 oxidase which was constitutively-active under both aerobic and microaerobic conditions;
whereas, NADH peroxidase was positively-activated by the presence of oxygen and had
a long induction time in contrast to NADH oxidase. These observations indicated that
a coupled NADH oxidase - NADH peroxidase system was the main oxidative stress resistance
mechanism in L. panis PM1, and was regulated by oxygen availability. Under aerobic conditions, NADH is
mainly reoxidized by the NADH oxidase - peroxidase system rather than through the
production of ethanol (or 1,3-propanediol or succinic acid production if glycerol
or citric acid is available). This system helped L. panis PM1 directly use oxygen in its energy metabolism by producing extra ATP in contrast
to homofermentative lactobacilli.

Keywords:

Introduction

Lactobacillus panis PM1 is an aerotolerant and obligatory heterofermentative microorganism isolated from
bioethanol thin stillage, and has been the focus of attention due to its ability to
produce 1,3-propanediol (1,3-PDO) during the fermentation of glycerol under anaerobic
conditions (Khan et al. 2013). Lactobacillus panis belongs to the group III heterofermentative lactobacilli, which includes L. brevis, L. buchneri and L. reuteri, where the 6-phosphogluconate/phosphoketolase (6-PG/PK) pathway is the primary carbohydrate
fermentation pathway (Khan et al. 2013; Luthi-Peng et al. 2002; Pedersen et al. 2004; Veiga-da-Cunha and Foster 1992). In theory, when one glucose molecule is consumed, three NADH and one ATP molecules
are generated. Subsequently, one pyruvic acid and one acetyl phosphate molecules accept
protons from one and two NADH molecules, respectively, and regenerate NAD+. End-products of this metabolism are lactic acid and ethanol, respectively. Overall
heterolactic fermentation of glucose through the 6-PG/PK pathway results in 1 mol
each of lactic acid, ethanol, and CO2 and 1 mol ATP per mol glucose consumed (Kandler 1983).

For heterofermentative lactic acid bacteria (LAB), external electron acceptors can
be used as alternate routes to reproduce NAD+. The presence or absence of electron acceptors determine whether ethanol (and no
more ATP) or acetic acid (and 1 additional ATP) is produced from a glucose molecule
(Chen and McFeeters 1986; Condon 1987; McFeeters and Chen 1986; Talarico et al. 1990; Veiga-da-Cunha and Foster 1992). For example, when glycerol exists, the regeneration of NAD+ for glucose metabolism can be achieved through the conversion of glycerol to 1,3-PDO
using glycerol as the electron receptor (Saxena et al. 2009; Veiga-da-Cunha and Foster 1992). The presence of external electron acceptors, therefore, affects the energy metabolism
and end-product profiles, as well as further fermentation applications of LAB.

Therefore, the presence of oxygen in the growth environment of LAB will induce oxidative
stress to which bacteria have various responses mechanisms. A common oxidative stress
resistance mechanism found in LAB is a coupled NADH oxidase - NADH peroxidase system
(Miyoshi et al. 2003). In these coupled reactions, intracellular oxygen is first used to oxidize NADH
into NAD+ by NADH oxidase, thereby releasing H2O2. Subsequently, H2O2 is reduced to H2O by NADH peroxidase (Condon 1987; Higuchi et al. 2000; Miyoshi et al. 2003). However, the activity of NADH peroxidase is generally low (10 to 30 times lower
than that of NADH oxidase) in L. lactis and has not been detected in some latobacillus strains. Thus, cellular H2O2 detoxification is inefficient in some LAB capable of producing H2O2 under aerobic conditions (Anders et al. 1970; Komagata 1996).

Our previous research showed that the presence of oxygen during the fermentation of
glycerol by L. panis PM1 negatively affected cell growth, glucose consumption, and end-product production,
including 1,3-PDO. The protection mechanism towards oxidative stress is a key element
to optimize L. panis PM1 for 1,3-PDO production in biofuel waste material applications. The NADH oxidase
- NADH peroxidase system and conversion of glycerol to 1,3-PDO both use NADH as a
key factor for their reactions. Therefore, the clarification of the control of oxidative
stress by this strain can shed light on how it regulates 1,3-PDO production. In this
study, we clearly demonstrated the oxygen-dependent function of NADH oxidase and NADH
peroxidase and its involvement in the NAD+ regeneration system of L. panis PM1.

Materials and methods

Chemicals

All chemicals used in this study were ACS grade, or better, and purchased from Sigma-Aldrich
(St. Louis, MO, USA).

Bacterial strains and growth conditions

Lactobacillus panis PM1 was isolated from bioethanol thin stillage in our lab (International Depository
Authority of Canada; accession number 180310–01). Strain PM1 was cultured at 37°C
using commercial MRS medium (BD, Franklin Lakes, NJ, USA) until late log phase and
was then transferred to modified MRS medium (mMRS). The mMRS medium consisted of 10
g glucose, 5 g yeast extract, 10 g peptone, 10 g meat extract, 2 g K2HPO4, 2 g ammonium citrate, 5 g sodium acetate, 100 mg MgSO4·7H2O, 50 mg MnSO4, along with a defined concentration of electron acceptors, such as citric acid (26
mM) or glycerol (160 mM) per litre. The cultures were incubated at 37°C under aerobic
or microaerobic conditions. Aerobic and microaerobic cultures were grown using the
same medium and temperature. Continuous aeration was provided to aerobic cultures
by agitation; whereas, air-tight 15 ml tubes, filled to the two-thirds level, were
incubated under static conditions to establish microaerobic conditions. It should
be noted that our previous study (Khan et al. 2013) indicated there was little difference in the behaviour of L. panis PM1 under anaerobic and microaerobic conditions, thus we did not include anaerobic
culture in this study.

Quantification of H2O2 production

Lactobacillus panis PM1 cells were removed from the culture media using centrifugation (14,000 × g, 5 min). Hydrogen peroxide concentrations of the cell-free media were measured in
accordance with the Pierce Quantitative Peroxide Assay Kit (Thermo Scientific, Rockford,
IL, USA) based on oxidation of ferrous to ferric ion in the presence of xylenol orange.

RNA preparation

RNA was extracted from Lactobacillus panis PM1 cells by the hot phenol extraction method, as described by Oh and So (2003) with minor modifications. Briefly, 10 ml of exponentially-growing bacteria from
liquid media were added to a tube containing 1.25 ml of ice-cold ethanol/phenol stop
solution (5% water-saturated phenol, pH < 4.5, in 95% ethanol), and harvested by centrifugation
for 5 min at 10,000 x g. The cell pellets were resuspended in 600 μl of diethylpyrocarbonate
(DEPC)-treated water. Glass beads (0.8 g, 452–600 μm in diameter; Sigma) and 600 μl
of pre-warmed acid-hot phenol:chloroform:isoamylacohol (PCI, 25:24:1, v/v) were added
to the cell suspensions, and the mixture was incubated at 65°C for 10 min with vigorous
vortexing for 30 sec duration every 30 sec. The samples were centrifuged for 10 min
at 14,000 x g and then the supernatants (500 μl) were transferred to fresh 1.5-ml
micro-tubes containing 500 μl of the pre-warmed PCI and incubated at 65°C for 5 min
with vortexing every 30 sec. Samples were then centrifuged for 10 min at 14,000 x
g. The aqueous layer (400 μl) was transferred into new 1.5-ml micro-tubes and mixed
with 95% ethanol (800 μl) and 3 M sodium acetate (40 μl). The mixtures were kept at
−80°C for 30 min and centrifuged at 14,000 x g. The RNA pellet was washed with 70%
ethanol and resuspended in 50 μl of RNase-free water. RNA was treated with DNase and
purified using the RNeasy kit (Qiagen, Toronto, ON, Canada). The quantity of RNA was
determined by measuring the absorbance at 260 nm (optical density (OD) 1 at A260 = 40 μg · ml-1 RNA), using a DU 800 spectrophotometer (Beckman Coulter, Mississauga, ON, Canada),
and its purity was determined by measuring the A260/A280 ratio.

Reverse transcription

The primers used in this study were specifically designed by Primer3 (http://frodo.wi.mit.edu/primer3/webcite) for real time RT-PCR applications of L. panis PM1 from the nucleotide sequence of the annotation data (Table 1). The RT reaction mixture contained 0.5 μg of total RNA and 0.25 μM of reverse primers
of the selected genes. The RT reaction was performed using qScript cDNA SuperMix (Quanta
Biosciences, Inc., Gaithersburg, MD, USA) according to the manufacturer’s instructions.
The reaction mixture was incubated at 25°C for 5 min and at 42°C for 30 min, and the
reaction was terminated by incubation at 85°C for 5 min with a Techne thermal cycler
(FTGENE-5D, Techgene, Burlington, NJ, USA).

Quantitative PCR

Real-time PCR amplification was performed in a CFX96 real-time detection system (Bio-Rad,
Hercules, CA, USA) using SsoFast EvaGreen Supermix (Bio-Rad). The total volume of
the PCR master mixture was 20 μl, to which cDNA template equivalent to 25 ng RNA starting
material and 0.5 μM of each primer (Table 1) was added. PCR amplification was initiated at 95°C for 30 s followed by 40 cycles
of 95°C for 5 s and 60°C for 10 s. Amplification was followed by a melt-curve analysis
between 65°C and 95°C using a 0.5°C increment. All sample and primer combinations
were assessed in three biological replicates with two technical replicates per biological
replicate. A no-template control was used for the negative control PCR, and PCR specificity
and product detection were verified by examining the temperature-dependent melting
curves of the PCR products and ethidium bromide staining on 1% agarose gel. For relative
gene expression, the 2-ΔΔCt method using the 16S rRNA gene as the normalizer was performed as described by Livak
and Schmittgen (2001). The steps for calculating the expression ratio are following:

Normalized expression ratio of nox and npx(test) = 2–ΔΔCt.

The Real-Time data were processed using CFX Manager Software (Bio-Rad).

Preparation of crude extracts

Lactobacillus panis PM1 cells grown to mid-exponential phase under microaerobic or aerobic conditions
were harvested by centrifugation, washed with 100 mM phosphate buffer (pH 7.0), and
the cells in pellets were then disrupted using sonication (three times for 1 min with
a 3 min rest interval at output level 2, Sonifier 450; Branson, CT, USA) using the
same buffer. Crude extract was obtained by centrifugation for 10 min at 14,000 x g,
and protein concentration was determined using the Protein Assay Kit (Bio-Rad) with
bovine serum albumin (BSA) as a standard.

Enzyme assay

NADH oxidase and NADH peroxidase activities were determined by measuring the H2O2 concentration generated and decomposed by the crude extracts, respectively. The assay
mixture contained 200 μM of the reduced form of nicotinamide adenine dinucleotide
(NADH) and 20 μM flavin adenine dinucleotide (FAD+) in 50 mM phosphate buffer at pH 6.0. The assay was carried out at 30°C under aerated
or microaerobic conditions. For the NADH peroxidase assay, hydrogen peroxide was added
into the above assay mixture to an initial concentration of 30 μM. The concentrations
of H2O2 generated or decomposed were quantified as described above. In these determinations,
one unit of activity corresponds to the generation (for NADH oxidase) and decomposition
(for NADH peroxidase) of one μmol of H2O2 in one minute.

Determination of glucose and end-products

Culture optical density was measured as an index of growth at 600 nm with a DU 800
spectrophotometer. After centrifugation, the supernatant was filtered through 0.22-μm
pore size filters and stored at −20°C for HPLC analysis. To quantify the concentration
of glucose, organic acids and ethanol, samples were analyzed on an organic acid column
(HPX-87H; Bio-Rad) using an HPLC system equipped with a refractive index detector
(RID G1362A, 1100 series; Agilent Technologies, Palo Alto, CA, USA). Operating conditions
were determined by the method described in the column manual with minor modifications.
Filtered culture medium (40 μl) was loaded on the column and eluted with 5 mM sulfuric
acid at a flow rate of 0.6 ml/min at 55°C for 30 min.

Statistical analysis

For growth experiments and determinations of H2O2 concentrations, data are presented as the mean values calculated from at least three
independent experiments. For activities of NADH oxidase and NADH peroxidase, standard
errors of the means from at least three independent experiments were also calculated
and presented. Differences in culture and enzyme assay conditions with NADH oxidase
or NADH peroxidase activity (unit/mg protein) were analyzed by the t test (Mann–Whitney test) for two groups or the one-way ANOVA test (Kruskal-Wallis
test) for three groups using GraphPad Prism 5.0 software (GraphPad Software, Inc.,
San Diego, CA, USA). P < 0.05 was considered significant.

Results

Influence of oxygen on the physiology of L. panis PM1

The rates of growth during the first 24 hours of culture were similar between aerobic
and microaerobic L. panis PM1; however, the aerobically cultured L. panis PM1 entered stationary phase earlier than the microaerobic culture (Figures 1a and b). This early entry into stationary phase was associated with a halt in production
of end-products, but not with glucose depletion (as approximately 30 mM glucose remained
after 24 hours) (Figure 1a), indicating that glucose concentration was not a critical cause of the cellular
growth interruption. Unlike aerobically cultured samples, the microaerobic cultures
consumed all available glucose (55 mM) within 48 hours and produced nearly-equimolar
amounts of lactic acid and ethanol (Figure 1b), revealing a typical heterolactic fermentation of glucose through the 6-PG/PK pathway.
Also, the cells grown under microaerobic conditions were observed to consume lactic
acid during stationary phase, reducing the concentration of lactate from 53 mM to
33 mM. In contrast, aerobic cultures did not utilize lactic acid after cessation of
glucose consumption (Figure 1a). Furthermore, the ratio of ethanol production to glucose consumption (11:26 mM)
was less during aerobic culture than during microaerobic culture (55:55 mM). These
results indicated that, under aerobic conditions, an alternate metabolic route re-oxidized
NADH through the 6-PG/PK pathway while not forming ethanol.

Figure 1.Effect of oxygen and time on growth and end-product formation in L. panis PM1. Growth response (OD600) and end-product formation of L. panis PM1 cultivated in mMRS under aerobic (a) and microaerobic (b) conditions. Samples for cell density and HPLC analysis were withdrawn from the cultures
after 24, 48, 72 and 96 hours.

Production of H2O2 by aerobic culture and its effects on bacterial growth

Aerobic culture resulted in the production of ten-fold higher concentrations of H2O2 than during microaerobic culture. Rapid accumulation of H2O2, reaching approximate 100 μM, was achieved in the first 24 hours of aerobic culture
(Figure 2a). The concentration of H2O2 necessary to completely inhibit the growth of L. panis PM1 was determined to be approximately 120 μM H2O2 (Figure 2b). Accumulation of H2O2 reached close to this inhibitory concentration level within 24 hours of aerobic culture.
Concurrently, a reduction of cell density was observed after 24 hours aerobic culture
(Figure 1b). This data, therefore, indicated a clear association between H2O2 produced under aerobic conditions and the early entrance of L. panis PM1 into stationary phase.

Figure 2.Effect of oxygen and time on the accumulation of hydrogen peroxide and growth by
L. panis. Growth response (OD600) and H2O2 production by L. panis PM1 cultured in mMRS under aerobic (circles) and microaerobic (triangles) conditions
after 24, 48, and 72 hours (a). The inhibitory concentrations of H2O2 were determined from L. panis PM1 cultures grown in MRS broth containing H2O2 at concentration of 0, 30, 60, 120, 240, and 480 μM for 2 days under microaerobic
conditions. Optical density was measured at 600 nm with a spectrophotometer (b).

Specific activities of NADH oxidase and NADH peroxidase

The whole genome data of L. panis PM1 (unpublished draft data) revealed only candidate genes for NADH oxidase and NADH
peroxidase; whereas, other protective enzyme genes that might respond to the toxic
effects caused by oxygen were not detected. The expression levels of these two genes
were compared under aerobic and microaerobic conditions by qRT-PCR. It was determined
that nox (NADH oxidase gene) and npx (NADH peroxidase gene) were expressed at similar levels under both culture conditions
(Table 2) even though L. panis PM1 was shown to produce lethal levels of hydrogen peroxide under aerobic, but not
microaerobic, conditions. The levels of activity of NADH oxidase and NADH peroxidase
were measured from the cells grown under microaerobic and aerobic culture conditions.
The specific activity of NADH oxidase was comparable (P > 0.1) under both aerobic and microaerobic cultures (Table 2). Interestingly, the activities of NADH oxidase were dependent on availability of
oxygen in the respective assay reactions. When the specific activities of NADH oxidase
were compared between aerated and non-aerated assay conditions, significant differences
were observed (P < 0.05). Higher activities of NADH oxidase were observed in aerated assay than non-aerated
assay for both aerobically- and microaerobically-grown cultures (158.8 ± 7.6 vs. 92.5 ± 2.2
and 144.0 ± 2.0 vs. 103.1 ± 5.6 units/mg, respectively). In contrast to NADH oxidase,
NADH peroxidase activity was only detected in aerobic cultures. Enzyme assay conditions
significantly (P < 0.05) affected the levels of activity of NADH peroxidase in the opposite direction
of NADH oxidase; higher enzyme activity was observed under non-aerated assay conditions
(148.3 ± 9.7 vs. 197.3 ± 1.7 units/mg) (Table 2).

Role of oxygen in oxidative stress

Oxygen availability in the culture media directly affected the coupled NADH oxidase
- NADH peroxidase system of L. panis PM1, controlling the accumulation of H2O2 under aerobic conditions. When L. panis PM1 was cultured in 15-ml conical tubes containing 9, 6, and 3 ml mMRS under aerobic
conditions (in order to incrementally-increase oxygen availability in the aerobic
cultures), the H2O2 accumulation was greatest and most rapid in the 3-ml culture, reaching a maximal
value by 12 hours in all samples (Figure 3). The H2O2 concentration decreased after 12 hours in all samples; however, the degree of H2O2 decomposition occurred in proportion to oxygen availability in the culture media
(63% in the 3-ml culture, 33% in the 6-ml culture, and 13% in the 9-ml culture). The
final amount of cell growth was in accordance with the amount of H2O2 accumulated in the culture media until 12 hours (Table 3 and Figure 3). NADH oxidase activity was constitutively-expressed during the early stages of cell
culture, and high NADH oxidase activities were determined in the all 6-hour aerobic
cultures (approximately 100 units/mg). The activity of this enzyme increased in a
time-dependent manner until after 24 hours of culture. The specific activities of
NADH oxidase did not show significant differences in the three aerobic culture conditions
(P > 0.1). In contrast to NADH oxidase, NADH peroxidase specific activity was only observed
in the 3-ml 24-hour culture (Table 3). This data clearly demonstrated that NADH peroxidase activity was induced according
to oxygen availability that also elevated the production of H2O2 by NADH oxidase.

Table 3.The result of cell growth and specific activities of NADH oxidase and NADH peroxidase
according to oxygen availability

The change of NADH flux by NADH oxidase

Oxygen was a preferred electron acceptor for glycerol or citric acid and changed the
flux of NADH for reoxidation in L. panis PM1. Inhibitory levels of H2O2 were accumulated following 24 hour culture of L. panis PM1 in mMRS containing either citric acid (24 mM) or glycerol (160 mM) as electron
acceptors under aerobic conditions (124 and 120 μM H2O2, respectively). The consumption of glucose (11 and 27 mM in citric acid and glycerol
media, respectively) and production of ethanol (4 and 6 mM in citric acid and glycerol
media, respectively) were suppressed similar to that observed in aerobic cultures
lacking additional electron accepters (Figure 4). In addition, little citric acid (7 mM) or glycerol (13 mM) was consumed. The amount
of lactic acid produced correlated only with the amount of glucose utilized. Considering
the amount of citric acid or glycerol consumed and acetic acid produced (28 and 39
mM, respectively) in the culture media, it appeared that citric acid and glycerol
contributed only slightly to an increase in acetic acid production and utilization
for NADH recycling.

Discussion

We previously reported the aerotolerant nature of L. panis PM1 and its ability to use glycerol as the means of NADH recycling in the absence
of oxygen (Khan et al. 2013). However, the presence of oxygen prevented 1,3-PDO formation and thus markedly-affected
NADH recycling in this strain. In this study, the influence of oxygen on NADH recycling
system and the oxidative stress resistance mechanism in its aerotolerance was investigated.
Moreover, the metabolic profile was further investigated to understand how oxidative
stress resistance mechanisms of L. panis PM1 influenced the profile of metabolic end-products.

During aerobic culture, L. panis PM1 prematurely entered into a stationary phase without depleting glucose (Figure
1a). This early entry into stationary phase was also associated with a ten-fold higher
accumulation of H2O2 compared with microaerobic culture (Figure 2a). Therefore, the accumulation of H2O2 in aerobic culture was an apparent reason for the early cessation of growth. Anaerobic
metabolism theoretically makes one ethanol per every glucose consumed, but the presence
of oxygen altered this pattern to less than 1:1 ratio. These observations suggested
that H2O2 could be a main end-product of an alternate pathway for NADH recycling under aerobic
conditions, and that this could compete with NAD+-regeneration through ethanol production.

The production of H2O2 by LAB grown under aerobic conditions is commonly the result of flavoprotein oxidases,
including NADH oxidase, pyruvate oxidase, α-glycerophosphate oxidase, and superoxide
dismutase (Condon 1987). However, candidate genes for these enzymes were not found in the draft genome data
of L. panis PM1, with the exception of NADH oxidase. Pyruvate oxidase has been documented in
a few species of lactobacilli and is known to convert pyruvate to CO2 and acetyl phosphate, along with the formation H2O2 (Condon 1987). Pyruvate oxidase has its highest activity during the early stationary phase of
growth and is induced and repressed by oxygen and glucose, respectively, in L. plantarum (Saxena et al. 2009; Veiga-da-Cunha and Foster 1992). However, the presence of pyruvate oxidase does not adequately explain the early
entry into stationary phase observed during the aerobic culture of L. panis PM1. Our results showed that most of pyruvate produced during glucose consumption
was used to produce lactate in aerobic culture (Figure 1a), indicating that pyruvate oxidase apparently removed little pyruvate from this
pathway. NADH oxidase is the most common enzyme responsible for the production of
H2O2 from oxygen and is highly-active in LAB (Condon 1987; Higuchi et al. 2000; Tachon et al. 2011). LAB are known to possess either a NADH: H2O2 or a NADH: H2O oxidase, or sometimes both (Condon 1987; Higuchi et al. 2000). Final products of the reaction of NADH oxidase include either NAD+ and H2O2, or NAD+ and H2O, depending on whether two- or four-electrons are transferred by NADH: H2O2 oxidase or NADH: H2O oxidase (Condon 1987; Higuchi et al. 2000; Miyoshi et al. 2003). Our results showed that the crude extract from L. panis PM1 cultured under aerobic and microaerobic conditions could directly produce H2O2 using oxygen as a substrate, and the activity of the enzyme was found to increase
with the addition of FAD+ as well as aeration of the assay mixture (approximately 1.5 fold). These results
indicated that the NADH oxidase in L. panis PM1 was a NADH: H2O2 oxidase and a flavoprotein-like NADH oxidase, as seen in other gram-positive bacteria
(Komagata 1996; Marty-Teysset et al. 2000; Tachon et al. 2011).

Most LAB can respond (and protect themselves) to high concentrations of H2O2 produced through their oxidase enzymes during sugar fermentation (Higuchi et al.
2000). In fact, most LAB possess NADH peroxidase or pseudocatalase, and superoxide dismutase
exists in some LAB (Condon 1987). These enzymes can enable LAB to overcome otherwise-lethal concentrations of hydrogen
peroxide. The annotation data of the L. panis PM1 genome sequence and the results of the enzyme assays of NADH oxidase and NADH
peroxidase suggest that these enzymes are main factors in oxidative stress resistance.
The levels of accumulated H2O2 in the culture media could be accounted for by the differences in the activities
of NADH peoxidase and NADH oxidase. Our qRT-PCR analyses showed that oxygen did not
regulate nox and npx at the transcriptional-level, and mainly affected enzyme activities in L. panis PM1 (Table 2). While transcription levels were similar, activity assays exhibited that NADH peroxidase
was positively-activated by oxygen but required a long induction time to express activity
contrary to NADH oxidase. The oxygen-availability analyses indicated that higher oxygen
availability in the 3-ml culture could provide higher amounts of substrate (oxygen)
for NADH oxidase, resulting in greater accumulation of H2O2 in the first 12 hours. In the subsequent 12 hours, the accumulated H2O2 was decomposed by NADH peroxidase activity. The degree of degradation of H2O2 was dependent on NADH peroxidase activity, and the amount of activity was in proportion
with oxygen availability (Figure 3 and Table 3). Therefore, we concluded that a coupled NADH oxidase - NADH peroxidase system, regulated
by oxygen availability, was a key oxidative stress resistance mechanism in L. panis PM1.

Accumulation of H2O2 by NADH oxidase has been reported in group I homofermentative lactobacilli, like
L. delbrueckii, where approximate 97% of NADH was reoxidized by lactate dehydrogenase and NADH oxidase
accounted for only 3% of NADH reoxidation (Marty-Teysset et al. 2000). Thus, NADH recycling in group I LAB depends on a pyruvate supply from glycolysis,
rather than oxygen. Unlike homofermentative lactobacilli, the presence of electron
acceptors, such as oxygen, citric acid, or glycerol, directly influenced the flux
of NADH reoxidation in L. panis PM1. In our other studies, when L. panis PM1 was cultured in mMRS containing citric acid (24 mM) and glycerol (150 mM) under
microaerobic conditions, the major changes in end-product formation included a decrease
in ethanol, an increase in acetic acid, and the production of succinic acid (19 mM)
and 1,3-PDO (88 mM), respectively (unpublished data). The results of HPLC analyses
in the present study showed that aerobic conditions negatively-affected the production
of ethanol relative to glucose consumption, regardless of the presence of electron
acceptors (Figures 1a and 4). Also, when L. panis PM1 was cultured under aerobic conditions in mMRS containing citric acid and glycerol,
oxygen was used as the preferred electron acceptor, resulting in a shift of NADH flux
along with a significant decrease of the production of succinic acid (4 mM) and 1,3-PDO
(7 mM) (Figure 4). This data indicated that the activity of NADH oxidase was a key mechanism for the
reoxidation of NADH during growth in aerobic culture.

In addition to oxidative stress responses, NADH oxidase can also help L. panis PM1 use oxygen during energy metabolism, directly. That is, the shift of NADH recycling
with molecular oxygen redirected acetyl phosphate, which normally would be used to
produce ethanol, to the formation of acetic acid. This acetic acid production via
acetate kinase can stoichiometrically generate ATP (Condon 1987). Thus, O2-directed NADH recycling should be advantageous with respect to energy metabolism.
However, regeneration of NAD+ via NADH oxidase in L. panis PM1 led to overproduction of H2O2 with subsequent negative effects on growth and end-product formation. Our findings
indicate that varied oxygen availabilities of culture environments would greatly affect
energy metabolism as well as oxidative stress of L. panis PM1. The formation of 1,3-PDO is a main route for NADH reoxidation in the presence
of glycerol under anaerobic conditions; whereas, under aerobic conditions, NADH recycling
largely occurs through NADH oxidase activity. The present study indicates that energy
metabolism via the NADH oxidase system explains why L. panis PM1 fails to produce 1,3-PDO under aerobic conditions.

Competing interest

The authors declare that they have no competing interest.

Acknowledgements

The authors acknowledge the Saskatchewan Agriculture Development Fund and Agricultural
Bioproducts Innovation Program of Agriculture and Agri-Food Canada for supporting
this research.